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Frostig RD, editor. In Vivo Optical Imaging of Brain Function. 2nd edition. Boca Raton (FL): CRC Press; 2009.

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In Vivo Optical Imaging of Brain Function. 2nd edition.

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Chapter 2Two-Photon Functional Imaging of Neuronal Activity



For centuries scientists have been fascinated by the structure of the brain, all the more since the middle of the 19th century when advances in staining techniques enabled high-resolution light microscopic studies of brain cell morphology. Compared to studying cell morphology, it has been more difficult to experimentally investigate the dynamic nature of brain cells, including their morphological transformations, as well as the molecular and electrical signaling events underlying their specific functions. Still today, it remains challenging to study neural dynamics on the cellular level in intact brains of living animals. For many years, in vivo studies of brain cell dynamics relied solely on electrical recordings of neuronal activity, using either extracellular recordings of neuronal spike patterns or intracellular recordings of membrane potential dynamics. Strong light-scattering of neural tissue generally precluded optical imaging with cellular and subcellular resolution in the intact brain (with few exceptions in favorable cases [1]). A major breakthrough in 1990 was the invention of two-photon excited fluorescence laser scanning microscopy (TPLSM) [2], which enabled optical studies of brain cell morphology and function in vivo [3,4]. The success of TPLSM has been fostered by the parallel development of various novel staining techniques for in vivo labeling of brain cells with fluorescent markers. Expression of variants of fluorescent proteins by genetic means [5,6] complements the in vivo imaging capabilities of TPLSM particularly well. The combination of high-contrast fluorescence labeling of brain cells—neurons as well as glial cells—with high-resolution imaging in vivo has opened new research fields in neuroscience. It is now possible to watch brain cells “at work” so that neuroscientists can directly study cellular and molecular mechanisms underlying both normal brain function as well as brain diseases. Because of its great potential for gaining fundamental insights into how the brain develops, how it computes, and how it can adapt, in vivo two-photon imaging of brain cell dynamics has enormously expanded over the past decade, and it is still growing at a rapid pace.

This chapter provides an overview of two-photon imaging of brain cell dynamics in vivo. Because the principles of two-photon fluorescence excitation and TPLSM technology are treated in detail in Chapter 3, I focus here on the principles of dynamic measurements of single-cell and network activity using state-of-the-art labeling methods and various laser-scanning approaches. A key aspect of “activity” is that it always refers to processes evolving in the temporal domain, be it electrical or chemical signaling, turnover of proteins, or changes in cell morphology. Therefore, imaging modes capable of reading out the “dynamics” of living brain cells over a wide range of time scales will be particularly highlighted. In vivo calcium imaging studies—as the most prominent, current approach for functional measurements—will be summarized, and examples from mammalian brain imaging will be provided on both the single-cell as well as the population level. The chapter ends with an outlook discussing ongoing developments that are expected to become important research directions in the next years.


2.2.1. Two-Photon Microscopy

Two-photon microscopy is a laser-scanning technique, in which a laser beam is focused through a microscope objective down to a micrometer-sized light spot in order to excite fluorescent molecules. At each moment in time, laser-scanning microscopes typically collect fluorescence light from a single location in the sample. Spatial information is gathered by moving (“scanning”) the laser focus within the tissue. For example, raster-like, line-by-line scanning of a horizontal plane is a standard technique for acquiring two-dimensional (2D) images, and plane-by-plane acquisition of image stacks is a standard method to collect fluorescence data from a sample volume (Figure 2.1A). Exceptions from single-beam, laser-scanning microscopes are multifocal microscopes, in which fluorescence is generated simultaneously in multiple laser foci but then needs to be spatially resolved with a camera detector [7]. Two-photon microscopes based on multifocal excitation have been rarely applied in vivo so far; we will not considered them further here.

FIGURE 2.1. In vivo two-photon imaging of individual neurons and neuronal populations.


In vivo two-photon imaging of individual neurons and neuronal populations. (A) Schematic of a two-photon microscope setup for in vivo imaging in the brain. Beam control includes beam expansion and adjustment of laser intensity. Fluorescence can be detected (more...)

The key difference of TPLSM compared to confocal laser scanning microscopy is the physical process of light absorption, which is followed by fluorescence emission. In a two-photon absorption process, two photons are absorbed virtually simultaneously (within <1 fs). Because the two photons combine their energies to promote the molecular transition, their individual energies are relatively low. Hence, excitation light is of longer wavelength compared to single-photon absorption, typically in the near-infrared wavelength range (700–1000 nm). The use of near-infrared light is the first key advantage of two-photon excitation because longer-wavelength light is less scattered in biological tissue, enabling larger penetration depths. The second important feature is that the fluorescence signal depends nonlinearly on excitation light intensity (on the square of the intensity in the case of two-photon excitation). This nonlinear dependence (in defining TPLSM as a “nonlinear microscopy technique”) is highly beneficial because it confines fluorescence generation to the focus spot. As a consequence, at each point in time fluorescence photons are generated only locally, and they can be correctly assigned to their point of origin in three-dimensional (3D) space irrespective of whether they are scattered on their way out of the tissue to the detector. In TPLSM a confocal pinhole in front of the detector is not necessary to achieve optical sectioning; it should be omitted and as-many-as-possible fluorescence photons should be collected. In summary, the longer-wavelength excitation and the intrinsic optical sectioning property make TPLSM less sensitive to light scattering compared to confocal microscopy, clearing the view deep into the living tissue. For more details about the principles of two-photon fluorescence excitation, the reader is referred to Chapter 3 and a number of reviews [8–11].

2.2.2. Monitoring Cell Structure and Function

The special features of TPLSM make it ideally suited for the study of dynamic processes on the cellular and subcellular level in the intact nervous system. Dynamic events may comprise structural changes (i.e., morphological changes of individual cells or cell migration, and electrical and biochemical activity of cells, e.g., the generation of action potentials, ion fluxes across the membrane, or enzyme activities). Besides imaging neuronal dynamics TPLSM also has proved beneficial for measuring capillary blood flow [12] (see also Chapter 3) as well as activity in electrically nonexcitable glial cells (for review see References [13,14]). A key prerequisite for two-photon imaging is fluorescence labeling of the structures of interest with either an anatomical marker or a functional probe. Labeling everything generally is not very helpful; rather, a sparse, high-contrast label of particular cells is desirable. Meanwhile, several techniques for specific and sparse labeling are available, as will be discussed in Section 2.3.3.

Following the initial demonstration of the advantages of TPLSM for neuroscientific research [15], the first report on in vivo functional imaging of single neurons [4] was published in 1997. Figure 2.1B shows an image from a similar, more recent experiment. Here, an individual pyramidal neuron in the neocortex of an anesthetized rat was loaded with a fluorescent calcium indicator dye through a whole-cell patch pipette. This type of experiment has been useful for measuring calcium dynamics in neuronal dendrites and to study dendritic excitation [16–20]. Figure 2.1B in addition shows how a cell population can be easily visualized in the living brain using a simple negative-contrast stain. Negative stains can help to identify and to specifically target cells [21–23]. Note the difference between the standard xy-view as it typically appears during the experiment and the side view (or “xz-view,” with z being the dimension along the optical axis). Usually the side view is calculated offline from a stack of images; only recently sideways imaging has been implemented as a normal imaging mode in the microscope setup using a fast piezoelectric focusing device [24,25].

Early on it was noted that TPLSM also is a useful tool for precise photoma-nipulation [3]. This type of application is complementary to the imaging mode and has provided important results in brain slices during the last decade [26–28]. It still needs to be adapted for in vivo conditions perhaps fostered by the recent developments of light-gated cation channel proteins [29].

Two-photon microscopy covers the spatial scales between a micron and a millimeter. Its spatial resolution is good enough to resolve synaptic structures such as dendritic spines and axonal boutons in the intact brain. A number of studies have investigated structural dynamics of dendritic spines or axonal boutons [30–32], especially those modifying the connectivity scheme and thus probably contributing to neural circuit plasticity [33]. In the neocortex, structural dynamics has been found to be particularly high in microglial cells, the immunocompetent cells in the brain; in their “resting state” microglia continually expand and retract their fine processes, on a minutes time scale presumably subserving a homeostatic function [34,35]. Compared to these studies of morphological changes, functional measurements at the single-synapse level in vivo are more difficult; calcium signals in presynaptic terminals or dendritic spines have not been studied in vivo yet. An obvious difficulty is that remaining heartbeat and breathing-induced tissue pulsations make measurements remain on the micron scale difficult, even with good dampening.

On the other end of the spatial scales, the maximal imaging field of TPLSM covers larger and larger volumes. This expansion is driven by special high NA objectives with low magnification [36] and new labeling techniques for measuring activity patterns in distributed cell networks [37]. At present, the side length of typical field-of-views is on the order of a few hundred microns but may reach a millimeter soon. For the future, a major challenge is to cover even larger areas or 3D volumes while maintaining cellular resolution and sufficient temporal resolution. Most likely, such developments will require new technologies.

In summary, TPLSM is a versatile technique to image both structural dynamics and cell function in the living brain (Figure 2.2A). Despite the substantially reduced sensitivity of TPLSM to light scattering, two-photon imaging studies are still limited in terms of depth penetration. The maximal imaging depth that has been reached in the neocortex is about 1 mm, and background fluorescence generated during deep imaging sets a fundamental limit to the achievable depth [38,39]. TPLSM, therefore, is restricted to near-surface areas of the brain. So far it has been mainly applied to easily accessible brain areas such as neocortex [4], cerebellum [40], olfactory bulb [18], and, more recently, spinal cord dorsal horn [41]. Special procedures are required to reach deeper brain regions such as the hippocampus. Particularly promising are rodlike lenses (so-called gradient-index (GRIN) lenses), which have diameters below a millimeter and can be targeted to deeper areas for endoscopic imaging (Figure 2.2B). For example, using such an approach, fluorescently stained pyramidal cells have been visualized in the hippocampus of living mice [42,43]. In this chapter we will provide examples from the mammalian neocortex and the cerebellar cortex for illustration.

FIGURE 2.2. Overview of functional studies using two-photon microscopy.


Overview of functional studies using two-photon microscopy. (A) Most common read-outs are neuronal and glial morphological changes as well as measurements of intracellular calcium signaling. Another important application is the measurement of local blood (more...)


2.3.1. Two-Photon Microscope Setup

The principle setup of a two-photon microscope is similar to a confocal microscope, and in some aspects even simpler [9]. A main difference is the light source for fluorescence excitation. While wavelength-tunable Ti:sapphire IR lasers as sources of femtosecond laser pulses have developed into user-friendly key-turn instruments, they still are expensive and constitute a major cost for setting up a TPLSM. For several reasons, a number of research labs prefer to custom-build the core microscope part. Besides reducing the overall cost for a TPLSM system, this approach provides more flexibility regarding space issues (fitting an animal underneath the microscope) and scanning modes. New approaches for in vivo imaging often require creative solutions and microscope manufacturers tend to lag behind the rapid developments in the field. Because the technological aspects of setting up an in vivo TPLSM are extensively treated in a different chapter of this book (Chapter 3) I do not provide any further details on instrumentation here but rather focus on special laser-scanning modes and fluorescence labeling techniques.

2.3.2. Laser-Scanning Modes for Measurements on Various Timescales

A key technical aspect is the laser scanning technology. Following the invention of confocal laser scanning microscopy, most samples studied were fixed tissue sections with no need for fast dynamic imaging. For live cell imaging, in particular using TPLSM for imaging in the intact brain, the technical constraints of laser scanning are critical, and novel modes of laser scanning may be required. Meanwhile, a variety of scanning modes has been devised, enabling dynamic measurements of brain cell activity on a wide range of time scales (Figure 2.3). On the one end of the spectrum, simple line-scan technologies permit measurements with millisecond temporal resolution (Figure 2.3A,B). Because spatiotemporal patterns of neuronal spikes occur on the millisecond timescale the ability to image at high speed eventually will be crucial for characterizing large-scale activity patterns. Unfortunately, imaging of voltage-sensitive dyes—providing a direct measure of electrical activity—is still difficult to achieve with cellular resolution. As an alternative, fluorescent calcium indicators can indirectly probe spiking activity, although the achievable temporal resolution ultimately is limited by their calcium-binding kinetics. It thus remains a challenge to achieve in vivo measurements of large-scale population activity with high temporal accuracy using TPLSM.

FIGURE 2.3. Laser-scanning modes for imaging on a wide range of timescales.


Laser-scanning modes for imaging on a wide range of timescales. (A) Straight line scans provide measurements from specific cellular structures such as dendrites with millisecond resolution. (B) Free-line scans along closed curved trajectories can be used (more...)

Another challenge is to extend existing scan technologies to 3D, since neural circuits operate in 3D space. One approach, which will be exemplified further below, is to combine mechanical scan devices (e.g., galvanometric scan mirrors and a mechanical focusing element) to create 3D scan trajectories [44] (Figure 2.3C,D). Alternatively, nonmechanical scan devices, namely, acousto-optical deflectors (AODs), can be employed. These devices use sound-wave-generated diffraction gratings in crystals to deflect the laser beam, and they can achieve transition times of a few microseconds, enabling “jumps” from point to point [45–47]. Most recently, this approach has also been extended to 3D (Figure 2.3E). All these developments highlight that creative handling of scan technologies rather than adhering to standard raster-like scan patterns is beneficial for tuning data acquisition to the needs of a specific experiment, in particular when spatial and speed considerations need to be traded off against each other. While the scanning modes mentioned so far are suitable on timescales ranging from milliseconds to seconds to minutes, imaging modes on longer timescales are desirable for other applications. For example, morphological changes of brain cells in the adult brain—either under normal conditions or in models of brain disease or injury—may be slow but significant. Time-lapse imaging studies can create a movie of such dynamic changes by repeatedly imaging the same field-of-view over minutes to hours (Figure 2.3F). The experimenter needs to make a decision though how often (at what time intervals) the images shall be taken. Besides the rate of the expected changes relevant considerations here are the strength of photobleaching, how much illumination the tissue can tolerate, and how large data sets can be handled for storage and analysis.

A practical aspect special to in vivo experiments are motion artifacts induced by heartbeat and breathing. Tissue movements should be dampened as much as possible (e.g., by covering a craniotomy with agar and a coverglass.) If pulsations remain, heartbeat triggered image acquisition can help to stabilize imaging [34]. In addition, samples may drift over longer times; in particular, changes of the focal plane may occur. While lateral drifts can be corrected for rather easily by using image alignment routines, changes in focal plane cannot be corrected offline. A safe way to circumvent this problem is to always acquire small image stacks (10–20 planes), with the structures of interest in the center of the image stack (Figure 2.3G). Clearly, such repeated acquisition of image stacks, often referred to as “4D imaging,” slows down the measurements, but intervals of <1 min can be easily achieved and are sufficient in many cases. In addition, four-dimensional (4D) imaging can be applied on much longer timescales by repeatedly imaging a particular tissue volume in the same animal over days, weeks, or even months [30,31,48]. For finding the same structures again and again, blood vessels or other coarse landmarks can be utilized. Long-term imaging highlights the in vivo advantage of TPLSM, enabling previously impossible longitudinal studies of brain cell dynamics in the normal and diseased brain.

2.3.3. Traditional Dyes and Fluorescent Proteins

Ideally, in vivo TPLSM is applied to brains with prelabeled cells. In fact, cells contain autofluorescent molecules (e.g., nicotinamide adenine dinucleotide [NADH]), which can be imaged and used to determine a metabolic state [49]. In most cases, however, additional fluorescence labeling is necessary. Negative fluorescence staining has already been mentioned as a simple method to visualize cell bodies and identify at least some cell types [21–23]. For positive staining, a broad toolbox of fluorescent dyes [50] and a spectrum of in vivo labeling techniques are now available. Several classes of fluorescent labels can be distinguished. A first group comprises synthetic organic dyes, often referred to as “traditional” dyes, which need to be introduced into cells by physical or chemical means. Fluorescent proteins establish a second class; they are genetically encoded, and their expression can be induced in specific cell types. A third “mixed” is to attach a fluorophore to a protein of interest by tagging the protein with a specific short peptide sequence (for review see Reference [51]). Molecules in all these classes come in different colors with emission spectra ranging from blue to near-infrared. For example, an entire spectrum of fluorescent protein is available, which is continually growing [52]. Multicolor imaging will become increasingly important in the future because it permits simultaneous monitoring of dynamic processes in multiple tissue components, thus allowing a direct view at their interactions. The recent introduction of combinatorial expression of fluorescent proteins in “Brainbow” transgenic animals [53,54], resulting in nearly a hundred hues for easy distinction of individual cells, most likely is only the beginning of a new multicolor area of in vivo imaging.

Traditional dyes can be loaded into cells using a variety of methods, including uptake from the extracellular space, intracellular recordings, or electroporation. Often these methods lack cell type specificity. An exception is the red fluorescent dye sulforhodamine-101, a derivative of Texas Red, which for unknown reasons specifically labels astroglia in rodent neocortex [55]. A few drops of sulforhodamine-101 briefly applied to the exposed brain surface are sufficient for bright labeling. Combined with tail-vein injection of fluorescent dyes for labeling the microvasculature [12] astroglia staining with sulforhodamine-101 can be recommended as a simple test stain for evaluating and optimizing the in vivo performance of a TPLSM setup. Fluorescent protein expression can be achieved using local methods such as viral transfections [56,57] and targeted single-cell DNA electroporation [23,58] or more globally by generating transgenic animals. A major advantage is that cell type specificity can be achieved if a specific promoter is available. Transgenic animals with cell-specific expression of fluorescent proteins thus make the ideal situation of in vivo imaging of specifically prelabeled cells reality. A further boost of the generation of transgenic animals can be expected driven by the further improvements of activity-dependent fluorescent proteins.

2.3.4. Activity-Dependent Probes

The distinction between “anatomical” and “functional” markers—between markers for cell morphology or protein localization on the one hand and those for particular physiological parameters on the other—is a vague one. Often cell function manifests itself as structural dynamics, either of gross cell morphology or of intracellular rearrangement of molecular components. More precisely one may speak of “activity-dependent probes” in the case of dyes that change their fluorescent properties in response to a specific cellular activity (e.g., in response to ion influx or the activation of an enzyme). Voltage-sensitive fluorescent dyes and fluorescent ion indicators are examples of traditional activity-dependent probes. In addition, ongoing research continually produces new activity-dependent fluorescent proteins and improves their performance for specific purposes. Sometimes, however, an activity-dependent effect may need to be suppressed, as in the case of unwanted pH-sensitivity of some fluorescent proteins. Today, the most commonly applied activity-dependent probes are fluorescent calcium indicators, which will be discussed in more detail in the next section.


2.4.1. Calcium Indicator Types and Loading Methods

Intracellular binding of calcium ions to proteins is fundamental to a multitude of cell functions. In particular, considering the role of calcium ions in synaptic transmission, it is the basis for neuronal communication and information processing. Not surprisingly, therefore, fluorescent calcium indicators are the most advanced functional indicators currently applied in neuroscience. The first synthetic fluorescent dyes developed by Roger Tsien around 1980 were based on calcium-chelating molecules such as BAPTA [59]. Over the past decades, a wide palette of synthetic calcium indicators has been created, now comprising dyes with vastly different calcium-binding affinities and different spectral properties, to name just two of the most important dye properties. Different affinities are desirable because calcium concentration changes span a huge range from nanomolar level (e.g., in large cell compartments) to micromolar level (e.g., in highly localized submembraneous calcium domains). For in vivo TPLSM nonratiometric indicators that are single-photon excited around 488 nm (e.g., Calcium Green, Fluo-4, or Oregon Green BAPTA) have proved most suitable. The reason is that they are easily two-photon excited at wavelengths between 800–900 nm and can be loaded into neurons using several methods. But also traditional UV-excitable dyes such as Fura-2 can be employed for in vivo two-photon imaging [60]. Applications of red fluorescent calcium indicators (e.g., Rhod-2) on the other hand are still rare, mostly because far fewer indicator types are available and because labeling is more problematic. Rhod-2 AM, for example, preferentially loads glial cells in vivo, and has not been used for neuronal imaging. Generally synthetic calcium indicators show low photobleaching rates with two-photon excitation and have no obvious pharmacological side effects aside from adding calcium buffering capacity to the cell.

Depending on the research focus, one may want to label an individual cell or entire cell populations. Fortunately, there are now several rather simple methods available, with which one can “titrate” the number of labeled cells (Figure 2.4). In the first single-cell in vivo imaging study [4], individual neurons were filled with a calcium indicator through an intracellular recording electrode (Figure 2.4A). This approach results in high-contrast labeling of the cell and all its dendrites, and therefore is well suited to study subcellular signaling and dendritic integration (see below). Intracellular loading can be performed through either a sharp electrode or a whole-cell patch pipette [19]. The experimental procedure can be alleviated by visually guiding the pipette tip to particular neurons, using either fluorescence counterstains [61] or negative contrast [23]. Even simpler is the application of brief electrical pulses through a dye-filled micropipette, causing focal electroporation of individual cells [22] or local electroporation of small groups of neurons [62] (Figure 2.4B). This type of labeling should be helpful for investigating excitation flow through local neural circuits because it permits discrimination of individual cells and zoom in on synaptic structures. Finally, it has been a long-standing goal in neuroscience to achieve functional labeling of entire local populations of neurons with the aim to optically resolve neuronal network dynamics in vivo. A major breakthrough was achieved in 2003 in Arthur Konnerth’s group, when they successfully loaded cells in vivo using the membrane-permeable, acetoxymethyl(AM)-ester form of calcium indicator dyes [37]. AM-dyes easily penetrate the cell membrane, but once the ester groups are cleaved by endogenous esterases—reconstituting the original charged indicator molecule—they are trapped in the cytosol. Although AM-ester loading dates back to the early 1980s [63], only in 2003 Konnerth’s group demonstrated that direct injection of a bolus of dissolved AM-ester dye into the brain parenchyma leads to dye uptake in essentially all cells within a circumscribed volume (Figure 2.4C) (the only exception being microglial cells, for which it is unclear in how far they are unlabeled by AM-ester loading). Since then multicell bolus loading (MCBL) has become a widespread method for in vivo investigations of the functional organization of local circuits. Practical tips can be found in a detailed protocol of this method [64] and in reviews [65,66].

FIGURE 2.4. Methods for in vivo labeling of neurons with calcium indicator.


Methods for in vivo labeling of neurons with calcium indicator. (A) Individual neurons can be filled during an intracellular recording using a sharp electrode or a whole-cell patch pipette. (B) Electroporation through a glass pipette permits dye loading (more...)

Promising is also the application of genetically encoded calcium indicators (GECIs). The first calcium-sensitive fluorescent protein constructs were introduced more then 10 years ago [67]. Clever design strategies and continual improvements have led to advanced constructs that now seem ready for in vivo applications in mammalian brains (for reviews, see References [68]– [70]). As an example, a troponin-C-based fusion protein of two fluorescent proteins could be stably expressed in transgenic animals, and showed large calcium signals associated with pyramidal neuron activity both in brain slices and in vivo [71] (Figure 2.4D). Two more recent papers demonstrated improved in vivo imaging using GECIs [72,73]. Besides the generation of transgenic animals, all other means of driving protein expression can be utilized, including transfection with viral vectors and electroporation with DNA. GECIs can also be expressed in neuronal groups following in utero electroporation [72], which may allow labeling of particular cell types, depending on the timing of the electroporation procedure during the embryonal development. In view of the rapid advances in protein design, it is likely that GECIs will soon find widespread use for in vivo calcium imaging studies.

2.4.2. Inferring Neuronal Spiking Activity from Calcium Signals

Calcium indicators are utilized in different ways to investigate various aspects of cellular signaling. Two general applications are (1) the study of the calcium-dependence of particular processes (e.g., of transmitter release or synaptic plasticity) and (2) the indirect inference of electrical excitation and spiking activity from the fluorescence measurement. The second type of application is based on the close linkage between electrical excitation and calcium influx, mediated by voltage-dependent calcium channels. For example, each action potential in a neuron typically leads to opening of calcium channels in the somatic membrane and in at least part of the dendritic tree. With a sensitive calcium indicator, the brief surge of calcium ions can be detected as a rapid fluorescence change, which subsequently decays back to resting fluorescence, reflecting calcium removal from the cytosol (Figure 2.5B). The decay time constant of such a stereotyped “calcium transient” depends on calcium buffering and extrusion mechanisms, and typically is in the range of a few hundred millisecond for somata [75]. Thus, the instantaneous rate of neuronal spikes can be indirectly inferred from the fluorescence measurements [76], and in the best cases even single action potentials can be detected [77–79]. Clearly, the sensitivity will vary between cell types, and one should also keep in mind that calcium ions might enter the cytosol via other entry routes (e.g., by local release from intracellular stores). Furthermore, the temporal precision of determining spike times is limited by the temporal resolution of the laser scanning technique, which typically is slow compared to electrical recordings. Despite these drawbacks, the optical detection of spiking activity has a special appeal because it promises to enable in vivo measurements of nearly complete spiking patterns (including nonactive neurons) in local neural circuits, perhaps sampling populations up to several hundred cells [44]. Large-scale population calcium imaging complements existing electrophysiological methods and may help to reveal fundamental principles of network dynamics.

FIGURE 2.5. Examples of in vivo calcium imaging of dendritic excitation.


Examples of in vivo calcium imaging of dendritic excitation. (A) Spontaneous subthreshold membrane potential fluctuation in a neocortical pyramidal cell in an urethane-anesthetized rat revealed by a dendritic whole-cell recording (B). Fast calcium measurement (more...)


2.5.1. Imaging Individual Cells

Single neurons are sophisticated computing devices responsible for synaptic integration. Most dendrites possess nonlinear (“active”) properties, which in principle make them capable of generating local regenerative potentials. Although various modes of dendritic integration have been carefully investigated in brain slices, it is still largely unknown which modes are actually used in the living brain and under what specific conditions. In vivo calcium imaging may help to address this question under physiological conditions. As a first step, several studies have taken a closer look at action-potential-evoked calcium signals in dendrites of neocortical pyramidal neurons. Action potentials generated at the initial axon segment backpropagate into the dendritic tree, providing an “I am active” feedback signal to synapses, which is important for synaptic plasticity. In pyramidal neurons action potential backpropagation into the apical dendrite typically is decremental, meaning that the amplitude gradually declines with distance from the soma. Because calcium influx depends on action potential amplitude and duration, dendritic calcium signals indicate how strongly action potentials are attenuated and whether backpropagation might be modulated. Consistent with previous brain slice experiments, in vivo studies have confirmed that backpropagation of fast action potentials varies in different cell types, from nonattenuated in mitral cells of olfactory bulb [18] to decremental in neocortical pyramidal neurons [16,20] to strongly attenuated in cerebellar Purkinje neurons [25]. In the typical experiment, an individual neuron is filled with calcium indicator via a sharp electrode or a patch pipette, and action-potential-evoked calcium signals are measured at variable distances from the soma following dye loading. Figure 2.5 shows an example of a measurement from a neocortical layer 2/3 neuron. A line-scan measurement was performed near the main bifurcation to compare dendritic calcium transients during periods of spontaneous network activity (so-called Up states in the anesthetized animal) and periods of network silence. Calcium transients were larger during Up states, indicating that backpropagation of action potentials is boosted during spontaneous cortical activity [78]. A practical problem is that dendrites of principal neurons often are oriented perpendicular to the brain surface as, for example, the main apical dendrite of neocortical pyramidal neurons. Hence, the standard xy view provides a limited view, mainly of dendritic cross sections (see Figure 2.1B). This problem can be circumvented by adding a fast-focusing device, such as a piezoelectric objective mount, to the microscope setup. The additional z-scan can be used to image arbitrarily oriented planes in 3D, creating viewing angles on dendrites similar to brain slice experiments [25]. Moreover, 3D free line-scan modes can be tuned to record along individual dendrites or from groups of dendrites [25]. These new scanning modes should facilitate the characterization of spatiotemporal dendritic excitation profiles in the intact brain.

Arbitrary plane imaging is particularly advantageous for imaging in the cerebellar cortex because the orientation of the imaging plane can be adjusted to match the 2D plane of Purkinje cell dendritic trees (Figure 2.5B). This approach has enabled a direct view of climbing-fiber-evoked dendritic calcium transients [25] and promises to allow a more detailed investigation of the integration of parallel fiber and climbing-fiber inputs. In general, the question of whether, where, and when local dendritic spikes occur in the intact brain is now the focus of dendritic research. Large calcium action potentials have been reported in distal apical dendrites large layer 5 pyramidal neurons in vivo [17], but their significance for cortical association is still unclear. Layer 2/3 pyramidal neurons apparently have a weaker propensity to generate dendritic spikes [81]. However, the generation of local spikes requires clustered and synchronous synaptic inputs [26] and therefore may crucially depend on behavioral state. High-resolution imaging using TPLSM, eventually in awake animals (see below), will be the key technology to solve these questions.

2.5.2. Imaging Population Activity

Monitoring population activity is the second major application of in vivo TPLSM. The MCBL technique is well suited to label entire neuronal and astroglial networks (Figure 2.6A), making it possible to record from many, if not all, cells in a local area and to analyze the spatial and temporal response patterns. In vivo calcium imaging thus is an ideal tool to study local neuronal network dynamics. During recent years, it has been applied to various brain areas, including the visual cortex of cats and rodents [82–84] and rodent barrel cortex [76,77]. Obvious questions that can be addressed are in how far subassemblies of neurons represent particular stimulus features and how neuronal correlations are distributed throughout the network. A fruitful approach is to combine TPLSM with other imaging modalities. For example, intrinsic optical imaging (Chapter 9) can be used to first obtain a functional map of a particular brain area (e.g., the primary somatosensory cortex) and then target dye loading and two-photon imaging to an identified area [76,81]. Eventually, it would be extremely informative to combine measurements of network function with post hoc high-resolution anatomical analysis of the underlying network connectivity [83].

FIGURE 2.6. Example of calcium imaging of population activity.


Example of calcium imaging of population activity. (A) Time-lapse recording of spontaneous activity in neocortical layer 2 neurons following multicell bolus loading with Oregon Green BAPTA-1 AM. Example fluorescence traces from three neurons are depicted (more...)

One major limitation still is the number of cells that can be recorded from with sufficient temporal resolution. Recently, we have introduced 3D line scanning technology as one possible approach to extend measurements to larger neural networks (Figure 2.3C). Using a mechanically “swinging” objective, this method enables measurements from several hundred cells in 3D with a sampling rate of 10 Hz for the entire population (Figure 2.6B). There still is room for improvements of this purely mechanical approach, and I expect that within a few years 3D measurements at 10 Hz or faster rates will be possible from a thousand or more cells. An alternative approach to this problem is the use of nonmechanical scanners, namely, acousto-optical deflectors (AODs). These devices use sound-wave-generated diffraction gratings in a crystal to deflect the laser beam, and they can achieve point-to-point transition times of a few microseconds [45–47]. Recently, it could be demonstrated that special arrangements of AODs can provide not only fast 2D but also 3D random access scanning by rapidly changing the divergence of the laser beam, which effectively shifts the focal plane (Figure 2.3E) [45]. Though currently restricted to rather small volumes, this approach may eventually enable optical imaging of network activity with millisecond time resolution.


The scope of this chapter was to provide an overview of the new possibilities opened by TPLSM to study brain cell dynamics in vivo. Within only one decade, in vivo TPLSM has proved extremely valuable for addressing fundamental questions of neuronal wiring and single-cell as well as network computation. Although these questions are far from being solved it is clear that, in vivo, TPLSM will be an indispensable tool for their further investigation. Spanning the range from the single synapses to local networks TPLSM builds a bridge between traditional electrophysiological methods, probing the activity in individual neurons or dispersed networks, and larger-scale measurements of brain dynamics (without cellular resolution) such as electroencephalography, fMRI, intrinsic optical imaging (Chapter 9), or voltage-sensitive dye imaging (Chapter 6). Because of its special advantages for deep tissue imaging TPLSM has also expanded in various other research fields beyond neuroscience, for example, to the study of cell dynamics in the immune system, in the kidney, or in tumors.

For the near future, I see several important research fields that are likely to benefit from in vivo two-photon imaging: (1) Repeated imaging of cellular or network activity should enable studies of the postnatal development of neural circuit activity. This goal will require repeated imaging sessions with activity-dependent probes over days and weeks [72], similar to the approach routinely used for structural imaging. Stable expression of genetically encoded indicators should make this type of measurement feasible. (2) The same approach might also enable in vivo studies of circuit plasticity. The challenge is to perform repeated imaging of the same cellular network (e.g., before and after a sensory deprivation protocol or following brain injury). Again such experiments most likely will rely on genetically encoded markers. Analysis of changes in sensory response probabilities or neuronal cross correlations may then reveal mechanisms underlying circuit plasticity. Data analysis will not be trivial, though, because of the high-dimensionality of functional data sets from large populations. (3) The application of TPLSM to study animal models of brain disease such as Alzheimer’s disease [86] or stroke [87] will further expand. Here, the ability to monitor dynamic processes of interacting cell types using multicolor imaging—for example, glial cells and neurons—will be especially important. While most of traditional neuropathology is based on looking for changes in morphology, TPLSM opens the avenue for investigating alterations in cell activity or network dynamics and for probing the effect of potential therapeutic interventions [88]. All these fields will also benefit from further developments of techniques for optical stimulation (or silencing), based on photomanipulation using caged compounds [26–28] or light-activated channels [86].

Finally, to establish the behavioral relevance of cellular and local network signaling, a key challenge is to perform in vivo imaging experiments in awake rather than anesthetized animals [90]. Two methodological routes toward this goal are possible (Figure 2.7). First, TPLSM can be applied to head-restrained animals that are trained to tolerate head-fixation. This has recently been demonstrated for imaging of cortical network activity following bulk loading of calcium indicators [91]. Although they were head-restrained, mice in this study actually were mobile as they could run on an air-supported styrofoam ball (Figure 2.7A). The advantage of the head-restrained approach is that a standard microscope can be used providing optimal imaging conditions. In addition, the special 3D scanning modes for measuring large-scale network activity could possibly be applied to behaving animals. A special problem to be solved is that motion artifacts are more pronounced under awake conditions. Motion correction algorithms will be crucial to correct spatial distortions in the fluorescence measurements as far as possible [91,92].

FIGURE 2.7. Routes toward high-resolution two-photon imaging of neural activity in awake behaving animals.


Routes toward high-resolution two-photon imaging of neural activity in awake behaving animals. (A) Imaging in head-restrained awake mice. The animal is mobile on an air-supported styrofoam ball. (B) Imaging in freely behaving animals using a head-mounted (more...)

The second promising method for high-resolution optical imaging in behaving animals is miniaturization of the microscope so that it can be carried by rats or even mice (Figure 2.7B). The principal idea is to uncouple a small microscope headpiece from all large experimental equipment by guiding the excitation light through an optical fiber. The small headpiece may contain lenses, a small scan device, and a photodetector. Fluorescence light can also be collected through an additional optical fiber and detected remotely. In 2001, imaging of blood vessels in the neocortex of freely moving rats was demonstrated with the first two-photon fiberscope [93], weighing about 20 g. Since then, various designs of miniaturized two-photon microscopes have been published (for a review see Reference [94]). Most recently, we introduced an ultracompact fiberscope design (<1 g) and applied it to imaging calcium signals in Purkinje cell dendrites in anesthetized rats [91]. Using a single photon fiberscope another study recently achieved optical recordings of Purkinje cell activity in freely moving mice [96]. The next major breakthrough—to optically record neural activity with cellular resolution in freely moving animals—is still pending. Both the head-restrained and the freely moving approach should be highly beneficial for relating dendritic signals as well as local network activity to the behavioral state and actual behavior of the animal. For example, these methods may allow proving the in vivo occurrence of local spikes in thin neuronal dendrites and to reveal their functional relevance. A direct correlation of neural activity with animal perception and behavior will be essential to uncover fundamental principles of information processing on the level of local circuits and to formulate models of how large-scale brain activity emerges from the lower-level brain cell dynamics.


Recent work in my laboratory was funded by the Max Planck Society, the University of Zurich, and grants from the Human Frontier Science Program and the Swiss National Science Foundation.


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